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Identification of Mycobacterial α-Glucan As
a Novel Ligand for DC-SIGN: Involvement of
Mycobacterial Capsular Polysaccharides in
Host Immune Modulation
Jeroen Geurtsen, Sunita Chedammi, Joram Mesters, Marlène
Cot, Nicole N. Driessen, Tounkang Sambou, Ryo Kakutani,
Roy Ummels, Janneke Maaskant, Hiroki Takata, Otto Baba,
Tatsuo Terashima, Nicolai Bovin, Christina M. J. E.
Vandenbroucke-Grauls, Jérôme Nigou, Germain Puzo, Anne
Lemassu, Mamadou Daffé and Ben J. Appelmelk
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The Journal of Immunology is published twice each month by
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Copyright © 2009 by The American Association of
Immunologists, Inc. All rights reserved.
Print ISSN: 0022-1767 Online ISSN: 1550-6606.
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J Immunol 2009; 183:5221-5231; Prepublished online 25
September 2009;
doi: 10.4049/jimmunol.0900768
http://www.jimmunol.org/content/183/8/5221
The Journal of Immunology
Identification of Mycobacterial ␣-Glucan As a Novel Ligand
for DC-SIGN: Involvement of Mycobacterial Capsular
Polysaccharides in Host Immune Modulation1
Jeroen Geurtsen,2* Sunita Chedammi,* Joram Mesters,* Marlène Cot,† Nicole N. Driessen,*
Tounkang Sambou,† Ryo Kakutani,‡ Roy Ummels,* Janneke Maaskant,* Hiroki Takata,‡
Otto Baba,§ Tatsuo Terashima,¶ Nicolai Bovin,储 Christina M. J. E. Vandenbroucke-Grauls,*
Jérôme Nigou,† Germain Puzo,† Anne Lemassu,† Mamadou Daffé,† and Ben J. Appelmelk*
T
uberculosis (TB),3 caused by the bacterium Mycobacterium tuberculosis, is a major cause of death worldwide
and kills ⬎1.7 million people per annum (1). Upon inhalation, M. tuberculosis infects alveolar macrophages in which it is
able to persist for extensive periods of time (2). Normally, the
infected macrophages are contained within so-called granulomas;
however, in a substantial number of cases (⬃10%), the bacterium
*Department of Medical Microbiology and Infection Control, VU University Medical
Center, Amsterdam, The Netherlands; †Centre National de la Recherche Scientifique,
Département Mécanismes Moléculaires des Infections Mycobacteriennes, Institut de
Pharmacologie et de Biologie Structurale, and Institut de Pharmacologie et de Biologie Structurale, Université de Toulouse, Université Paul Sabatier (Toulouse III), Toulouse, France; ‡Biochemical Research Laboratory, Ezaki Glico Co., Ltd, Nishiyodogawa-ku, Osaka, Japan, §Department of Biostructural Science and ¶Department of
Maxillofacial Anatomy, Tokyo Medical and Dental University, Tokyo, Japan; and
储
Carbohydrate Chemistry Laboratory, Shemyakin and Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, Moscow, Russia
Received for publication March 10, 2009. Accepted for publication August 6, 2009.
The costs of publication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked advertisement in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
1
This work was funded under the European Union Framework 6 program (ref.
37388): ImmunoVacTB: A new approach for developing a less immunosuppressive
tuberculosis vaccine.
2
Address correspondence and reprint requests to Dr. Jeroen Geurtsen, Department of
Medical Microbiology and Infection Control, VU University Medical Center, van der
Boechorststraat 7, 1081 BT Amsterdam, The Netherlands. E-mail address: jeroen.
[email protected]
3
Abbreviations used in this paper: TB, tuberculosis; BCG, bacillus Calmette-Guérin;
CR3, complement receptor 3; DC, dendritic cell; DC-SIGN, dendritic cell-specific
ICAM-3-grabbing nonintegrin; HSA, human serum albumin; ManLAM, mannosecapped lipoarabinomannan; MOI, multiplicity of infection; MR, mannose receptor;
NMR, nuclear magnetic resonance.
Copyright © 2009 by The American Association of Immunologists, Inc. 0022-1767/09/$2.00
www.jimmunol.org/cgi/doi/10.4049/jimmunol.0900768
escapes its containment and causes active disease (2, 3). The interaction between M. tuberculosis and the host immune system is
very complex (4, 5). An important question is how the bacillus
survives its hostile environment, that is, the intracellular compartments of the macrophage, and thereby is able to persist within its
host for many years. There are good indications that the distinctive
features of the mycobacterial cell envelope are of key importance
to this issue (6). Mycobacteria possess a unique cell envelope that
consists of three major entities: the plasma membrane, the cell
wall, and an outermost layer, known as the capsule (7). The plasma
membrane resembles that of other bacteria and consists of a symmetrical bilayer of phospholipids. The cell wall consists of two
segments: a lower segment of peptidoglycan covalently linked to a
heteropolysaccharide, the D-arabino-D-galactan, which itself is
linked to long chain fatty acids (C60–C90) called mycolic acids,
and an upper segment of free, intercalating glycolipids and waxes
(7). Recently, the mycolic acid layer, which is now known as the
“mycomembrane”, has been shown to be organized in a structure
that resembles the outer membrane of Gram-negative bacteria (8,
9). Compounds that are secreted across the mycomembrane are the
polysaccharides that make up the outermost layer of the cell-envelope, that is, the capsule (7). Bacterial capsules are protective
structures expressed by many pathogenic bacteria and have been
shown to be important for the successful colonization of the host
(10, 11). The mycobacterial capsule is loosely attached to the surface and is mainly composed of proteins and polysaccharides (7).
Part of the capsular material is released into the environment of the
mycobacteria and is found in the culture filtrate (12). The polysaccharide composition of the capsule is conserved among mycobacteria and predominantly consists of an ␣-D-(134)-glucosyl
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Mycobacterium tuberculosis possesses a variety of immunomodulatory factors that influence the host immune response. When the
bacillus encounters its target cell, the outermost components of its cell envelope are the first to interact. Mycobacteria, including
M. tuberculosis, are surrounded by a loosely attached capsule that is mainly composed of proteins and polysaccharides. Although
the chemical composition of the capsule is relatively well studied, its biological function is only poorly understood. The aim of this
study was to further assess the functional role of the mycobacterial capsule by identifying host receptors that recognize its
constituents. We focused on ␣-glucan, which is the dominant capsular polysaccharide. Here we demonstrate that M. tuberculosis
␣-glucan is a novel ligand for the C-type lectin DC-SIGN (dendritic cell-specific ICAM-3-grabbing nonintegrin). By using related
glycogen structures, we show that recognition of ␣-glucans by DC-SIGN is a general feature and that the interaction is mediated
by internal glucosyl residues. As for mannose-capped lipoarabinomannan, an abundant mycobacterial cell wall-associated glycolipid, binding of ␣-glucan to DC-SIGN stimulated the production of immunosuppressive IL-10 by LPS-activated monocytederived dendritic cells. By using specific inhibitors, we show that this IL-10 induction was DC-SIGN-dependent and also required
acetylation of NF-␬B. Finally, we demonstrate that purified M. tuberculosis ␣-glucan, in contrast to what has been reported for
fungal ␣-glucan, was unable to activate TLR2. The Journal of Immunology, 2009, 183: 5221–5231.
5222
MYCOBACTERIAL ␣-GLUCAN: A NOVEL LIGAND FOR DC-SIGN
Materials and Methods
Purification of ␣-glucan
␣-Glucan was isolated from M. tuberculosis strain H37Rv as previously
described (14). In short, M. tuberculosis H37Rv was grown on synthetic
Sauton’s medium (24) as surface pellicles at 37°C. After 3 wk, during
exponential growth phase, the culture medium was filtered through a
0.22-␮m filter (Nalgene) to remove intact bacteria, concentrated 10-fold
under vacuum, after which the macromolecules were precipitated by adding 6 vol of cold ethanol overnight at 4°C. The precipitate was collected by
centrifugation (1 h at 14,000 ⫻ g), dissolved in and dialyzed against distilled H2O, and lyophilized. The crude mixture was then dissolved in 0.01
M NH4Cl (pH 8.35) and loaded on a DEAE-Trisacryl column (27 ⫻ 1 cm).
Neutral polysaccharides were collected from the void volume, after which
␣-glucan was separated from other neutral polysaccharides by gel permeation chromatography using a Bio-Gel P-200 column (100 –200 mesh,
80 ⫻ 1 cm; Bio-Rad). After elution with 0.5% acetic acid, the fractions
were concentrated under vacuum and lyophilized. Purified ␣-glucan was
dissolved in double-distilled H2O and checked for purity using gas chro-
matography of trimethylsylilated monosaccharides liberated by trifluoroacetic acid hydrolysis and 1H nuclear magnetic resonance (NMR) analysis
(characteristic signals between 5.4 and 3.4 ppm). Mannose or arabinose
was not detected by any of these methods. Finally, ␣-glucan was treated in
batches with Affi-Prep polymyxin matrix (Bio-Rad) to remove possible
endotoxin contaminants. Removal of endotoxins was verified using the
Kinetic-QCL chromogenic Limulus amebocyte lysate assay (Lonza).
Treatment of ␣-glucan with amyloglucosidase, hydrogen
peroxide, and phenol
␣-Glucan was treated with amyloglucosidase (Sigma-Aldrich; A3514, EC
3.2.1.3) by solubilizing 2 mg of purified ␣-glucan in 500 ␮l of acetate buffer
(0.05 M, pH 4.5). After this, the suspension was separated into two equal
fractions and 20 ␮l of amyloglucosidase (566.4 U/ml) or buffer was added.
Samples were left overnight at 55°C, after which the reaction was stopped
by heating for 90 s at 100°C. Samples were deionized in batches using
TMD8 (Sigma-Aldrich; M8157) and lyophilized. Degradation was confirmed by 1H NMR; signals at 5.38 and 5.00 ppm, present in the control
sample and characteristic of ␣-D-(134)-glucosyl and ␣-D-(136)-glucosyl
linkages, completely disappeared after enzymatic treatment. ␣-Glucan and
Pam3CysK4 (InvivoGen) were treated with hydrogen peroxide as previously described (25, 26). In short, ␣-glucan (0.5 mg/ml) and Pam3CysK4
(1 ␮g/ml) were incubated for 48 h in the dark at 4°C in the absence or
presence of 1% hydrogen peroxide. After incubation, the samples were
snap-frozen using liquid nitrogen and subsequently lyophilized. For treatment with phenol, 2 mg of purified ␣-glucan was solubilized in 2 ml of
H2O, after which 2 ml of phenol was added for 1 h at 60°C. Then, the
mixture was centrifuged at 4°C for 10 min (1700 ⫻ g), after which the
aqueous phase was collected and resubmitted to the same treatment. Finally, the aqueous phase was dialyzed against double-distilled H2O and
lyophilized. In all cases, the lyophilized ␣-glucan was dissolved in pyrogen-free water and stored at ⫺20°C for further analysis.
Detection of ␣-glucan with a cross-reactive mAb directed
against glycogen
For detection of ␣-glucan with the mAb, serial dilutions of ␣-glucan were
spotted on a methanol-activated polyvinylidene fluoride membrane and
baked for 1 h at 70°C. Thereafter, the membrane was washed with PBS/
0.05% Tween 80, blocked with blocking solution (Boehringer Mannheim),
and probed with the mAb (27). Following incubation, the membrane was
washed with PBS/0.05% Tween 80, incubated with peroxidase-labeled
goat anti-mouse IgM (American Qualex), and developed using 3,3⬘-diaminobenzidine tetrahydrochloride and 4-chloronaphthol.
Human DC and macrophage generation and cell culture
Macrophages and immature human DCs were generated from human
PBMCs. In short, PBMCs were isolated from heparinized blood from
healthy volunteers (Sanquin Bloodbank, Amsterdam, The Netherlands) using density-gradient centrifugation over a Ficoll gradient (Amersham Biosciences). PBMC fractions were washed six times with 50 ml of cold PBS
containing 0.5% sodium citrate (w/v). Next, monocytes were isolated from
PBMCs by a CD14 selection step using the MidiMACS system (Miltenyi
Biotec) (CD14-depleted PBMCs were used as PBLs in the bead-binding
assay with primary immune cells (see Fig. 1)). Monocytes were differentiated into immature DCs or macrophages (type 1 and type 2) in RPMI
1640 medium supplemented with 10% FCS, 100 U/ml penicillin and 100
␮g/ml streptomycin (all from Invitrogen) in the presence of 500 U/ml
recombinant human IL-4 and 800 U/ml recombinant human GM-CSF for
generating DCs, or in the presence of 50 U/ml GM-CSF for generating
macrophage type 1, or in the presence of 50 ng/ml M-CSF for generating
macrophage type 2 (all from PeproTech) (28). For macrophage differentiation, fresh cytokines were added after 3 days of culture. At day 6, macrophages and immature DCs were harvested. DCs were positive for
CD11c, CD40, and DC-SIGN expression, negative for CD14 and CD83
expression, and they expressed low levels of CD80, CD86, TLR2, TLR4,
and HLA-DR as assessed by flow cytometry using FITC- or PE-labeled
Abs (eBioscience). Macrophages were positive for CD11c, CD14, and
CD40, negative for DC-SIGN and CD83, and they expressed low/intermediate levels of CD80, CD86, TLR2, TLR4, and HLA-DR. Raji cells and
Raji cells transfected with DC-SIGN (29) were maintained in RPMI 1640
medium supplemented with 10% FCS, 100 U/ml penicillin and 100 ␮g/ml
streptomycin. HEK293 cells transfected with TLR2 (30) were kept in
DMEM medium (Invitrogen) supplemented with 10% FCS, 100 U/ml penicillin and 100 ␮g/ml streptomycin, 0.5 mg/ml GW418 (Sigma-Aldrich), 2
mM L-glutamine, and 110 mg/L pyruvate. All cells were cultured at 37°C
in a 5% CO2 atmosphere.
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polymer, known as ␣-glucan (12, 13). ␣-Glucan production follows the growth curve of M. tuberculosis (13), and the structure of
the medium-released ␣-glucan is the same as ␣-glucan that is attached to the cell surface (12). The molecule has an apparent molecular mass of 1.3 ⫻ 107 and is expressed both in vitro and in vivo
(14 –17). Its structure resembles that of cytosolic glycogen and is
composed of repeating units of five or six ␣-D-(134)-glucosyl residues substituted at some 6-OH positions with an oligo-glucosyl side
chain (15, 16). Recently, a detailed structural comparison of ␣-glucan and glycogen, both derived from Mycobacterium bovis bacillus Calmette-Guérin (BCG), was performed (16). This analysis
showed that the two molecules are similar in terms of chemical
composition, degree of branching, and branching length. However,
the backbone chains of ␣-glucan are longer and its three-dimensional structure is more compact (16).
M. tuberculosis possesses a variety of immunomodulatory factors that influence the host immune response. When M. tuberculosis encounters its target cell, the outermost components of its cell
envelope are the first to interact. However, until now, most studies
have focused on the role of cell wall components in this process,
whereas the capsular constituents, that is, ␣-glucan, did not receive
much interest. In one of the few studies regarding the functional
role of the mycobacterial capsule, it was shown that in a number
of M. tuberculosis strains capsular polysaccharides mediate the
nonopsonic binding to complement receptor 3 (CR3) and inhibit
C3 surface deposition (18). It was postulated that CR3-mediated
uptake may promote intracellular survival by suppressing the production of IL-12 and prohibiting a respiratory burst (19, 20). Later
on, Stokes et al. showed that in some types of macrophages, capsular polysaccharides prevented phagocytosis, thereby possibly
promoting the uptake via CR3 (21). More recently, Gagliardi and
coworkers showed that mycobacterial capsular ␣-glucan blocked
CD1 expression and suppressed IL-12 production in monocytederived dendritic cells (DCs) (22). Finally, studies on M. bovis
BCG fractions to identify active components in its use as an immunotherapeutic agent against bladder cancer showed that ␣-glucan might be an active component (23). Although these studies
suggest that ␣-glucan fulfills an important biological function, the
underlying mechanism and host factors involved remain largely
unknown.
The aim of this study was to further assess the functional role of
the mycobacterial capsule by identifying ␣-glucan receptors
present on host immune cells. Herein we report that the C-type
lectin DC-SIGN (dendritic cell-specific ICAM-3-grabbing nonintegrin) represents an important host ␣-glucan receptor. Furthermore, our findings support the idea that DC-SIGN-␣-glucan interaction can modulate the effector functions of DCs.
The Journal of Immunology
Coating of fluorescent polystyrene beads
Purified ␣-glucan and biotin-labeled polyacrylamide neoglycoconjugates
(synthesized by Syntesome as described (31)) were coated onto 1.0-␮m
Fluoresbrite YG microspheres (Polysciences, catalog no. 17154) and
streptavidin Fluoresbrite YG microspheres (Polysciences, catalog no.
24161), respectively, by adding 50 ␮g of purified compound to 1 ml of
0.1 M carbonate-bicarbonate buffer (pH 9.6) containing 1.4 ⫻ 109 beads in
presiliconized tubes (Sigma-Aldrich). After overnight incubation at 4°C
(while gently rotating), the beads were collected by centrifugation and
incubated for 1 h in 1 ml of Tris-buffered saline supplemented with 1 mM
MgCl2 and 2 mM CaCl2 (TSM) containing 5% human serum albumin
(HSA) (Sigma-Aldrich). After this, the beads were washed four times with
1 ml of TSM/0.5% HSA and finally resuspended in this same buffer. The
bead concentration was determined by measuring the absorption of the
bead suspension and comparing it to the absorption of a serial dilution
series of beads with known concentrations at 441 nm. HSA control beads
were prepared in a similar manner, except that ␣-glucan was replaced
by H2O.
Cell binding assays
Lectin-Fc ELISA
␣-Glucan, bovine glycogen (Fluka, catalog no. 50571), or synthetic glycogen ␣1-ESG-A (33) (all at 1 ␮g/ml in saline (100 ␮l)) was coated on
Nunc Maxisorp plates (Roskilde) overnight at 4°C. Plates were blocked
with 1% HSA, after which DC-SIGN-Fc (34), dectin-1-Fc (35), or mannose receptor (MR) (carbohydrate recognition domains 4 –7)-Fc (36) (2
␮g/ml; all dissolved in TSM/0.05% Tween 80) was added for 2 h at room
temperature in the absence or presence of 5 mM EDTA, 50 ␮g/ml AZND2, or 2 mg/ml mannan. After incubation, the plates were washed four
times with TSM/0.05% Tween 80 and incubated with a goat anti-human
IgG Ab conjugated with peroxidase (Jackson ImmunoResearch Laboratories). After incubation, the plates were washed eight times with TSM/
0.05% Tween 80 and developed using o-phenylenediamine dihydrochloride. Absorption was measured at 490 nm using an EL808 Ultra microplate
reader (Bio-Tek Instruments).
Cell stimulation assays
Cells (HEK293 cells were first released by trypsinization) were washed
with and resuspended in culture medium at a concentration of 1.25 ⫻ 106
cells/ml. Eighty microliters of cell suspension (1 ⫻ 105 cells) was transferred to a sterile 96-well U-bottom plate (Greiner Bioscience) and left for
2 h, followed by incubation (in triplicate) with ␣-glucan in the absence or
presence of LPS (from Salmonella enterica serovar Abortusequi (SigmaAldrich L5886)), bovine glycogen, synthetic glycogen ␣1-ESG-A,
Pam3CysK4, or LPS alone (final stimulation volume of 100 ␮l). In some
cases, the cells were preincubated with DC-SIGN inhibitors 2 h before
stimulation (3 ␮M anacardic acid (Calbiochem), 20 ␮g/ml AZN-D2). Unstimulated cells served as controls in all experiments. Culture supernatants
were harvested after 24 h of incubation (37°C, 5% CO2) by centrifugation
and stored at ⫺80°C for cytokine measurements using an ELISA. In some
experiments, the cell pellets were pooled, washed three times with PBS,
and lysed for subsequent mRNA isolation and analysis by quantitative
real-time PCR.
mRNA isolation and quantitative real-time PCR
mRNA was isolated from DCs (stimulated as described above) using the
mRNA capture kit (Roche) and cDNA was synthesized with the SuperScript VILO cDNA synthesis kit (Invitrogen). For quantitative real-time
PCR analysis, PCR amplification was performed using Express SYBR
GreenER qPCR Universal Supermix (Invitrogen) in a LightCycler 480
(Roche). Analysis of mRNA expression was performed using specific
primers for IL-10 (GAGGCTACGGCGCTGTCAT (forward) and
CCACGGCCTTGCTCTTGTT (reverse)) and for GAPDH (CCATGTT
CGTCATGGGTGTG (forward) and GGTGCTAAGCAGTTGGTGGTG
(reverse) and Ct values were calculated using the second derivative maximum method. For each sample, the normalized amount of IL-10 mRNA
(Nt) was calculated using the following formula: Nt ⫽ 2(Ct(GAPDH)⫺Ct(IL-10))
(37). The relative IL-10 expression was calculated for each sample by
setting the Nt value for LPS-stimulated cells at 1.
Cytokine measurements using ELISA
Human IL-8, IL-10, and IL-12p40 concentrations in the supernatants of
stimulated cells were determined using ELISA according to the manufacturer’s instructions (Invitrogen).
p65 Phosphorylation and acetylation
DCs (2.5 ⫻ 105 cells) were stimulated for 1 h (37°C, 5% CO2) with ␣-glucan or mannose-capped lipoarabinomannan (ManLAM) (isolated from M.
tuberculosis and provided by J. Belisle, Colorado State University, as part
of the National Institutes of Health, National Institute of Allergy and Infectious Diseases Contract no. HHSN266200400091C, titled “Tuberculosis
Vaccine Testing and Research Materials” (complete standard operating
procedure can be found at www.cvmbs.colostate.edu/microbiology/tb/pdf/
lamlm.pdf)) in the absence or presence of LPS. DC nuclear extracts were
prepared with the NuncBuster protein extraction kit (Novagen). p65 from
nuclear extracts was captured with the Pathscan p65 sandwich ELISA kit
(Cell Signaling Technology). Specific p65 phosphorylation and acetylation
was detected with rabbit anti-phospho-p65 (Ser276) or anti-acetyl-p65
(Lys310) polyclonal Abs (all from Cell Signaling Technology).
Statistical analysis
Data were statistically analyzed using a Student’s t test (two-tailed, twosample equal variance). Differences were considered to be significant when
p ⬍ 0.05.
Results
Mycobacterial ␣-glucan interacts with human DCs
As a first step in identifying ␣-glucan receptors on host immune
cells, we performed a screen with different types of immune cells
for their ability to interact with ␣-glucan. For this, fluorescent
polystyrene beads were coated with ␣-glucan (purified from M.
tuberculosis strain H37Rv) and incubated with various types of
primary immune cells, including PBLs, monocytes, and monocytederived macrophages (type 1 and type 2) and DCs. After washing,
the percentage of fluorescently labeled cells was determined using
flow cytometry. As shown in Fig. 1, the ␣-glucan-coated beads, as
compared with the control beads coated with HSA, showed a significantly increased association with both types of macrophages,
albeit only at a high cell-to-bead ratio, and with the DCs. The
interaction with the cells was dose-dependent, as a higher beadto-cell ratio increased the association with the beads (Fig. 1). Although significant, the relative increase in the binding of the ␣-glucan-coated beads to the macrophages, as compared with the HSA
control beads, was only modest (macrophage type 1, 1.29 ⫾ 0.07;
macrophage type 2, 1.53 ⫾ 0.18) (Fig. 1). In contrast, the association of the ␣-glucan-coated beads with the DCs was more prominent and varied between a 2.08- to 2.86-fold increase as compared
with the control beads (Fig. 1). These results suggest that ␣-glucan
interacted with a receptor residing on the surface of the DCs.
Hence, we set out to identify this receptor.
Interaction of ␣-glucan with human DCs is dependent on
DC-SIGN
To investigate which lectin was responsible for the binding of
␣-glucan to DCs, a panel of lectin-Fc constructs representing DC
lectins known to interact with mycobacteria was tested for the
ability to bind to the molecule. As shown in Fig. 2A, ␣-glucan was
exclusively recognized by DC-SIGN-Fc but not by the other two
lectin-Fc constructs tested, that is, MR-Fc and dectin-1-Fc. DCSIGN is a Ca2⫹-dependent C-type lectin that was previously
shown to be the major mycobacterial receptor expressed by DCs
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Fluorescent beads or M. bovis BCG strain Copenhagen expressing dsRed
(grown in Middlebrook 7H9 broth (Difco) with 10% Middlebrook albumin/dextrose/catalase enrichment (BBL), 0.05% Tween 80, and 50 ␮g/ml
hygromycin) was added to cells (5 ⫻ 105 cells in 100 ␮l TSM/0.5% HSA)
and incubated for 30 min at 37°C in absence or presence of competitors (50
␮g/ml AZN-D2 (32), 2 mg/ml yeast mannan (Sigma-Aldrich), 50 ␮g/ml M.
tuberculosis ␣-glucan). AZN-D2 is specific for DC-SIGN and blocks ligand binding, whereas mannan occupies the DC-SIGN carbohydrate recognition domain and similarly blocks binding to other ligands. After washing, the percentage of fluorescent cells was determined using a FACScan
analytic flow cytometer (BD Biosciences) and analyzed using the manufacturer’s software (CellQuest version 3.1f).
5223
5224
MYCOBACTERIAL ␣-GLUCAN: A NOVEL LIGAND FOR DC-SIGN
FIGURE 1. Mycobacterial ␣-glucan interacts with human DCs. Primary human PBLs, monocytes, DC,
macrophages type 1 (M␾-1), and macrophages type 2 (M␾-2) were incubated with beads coated with ␣-glucan
or HSA for 30 min at 37°C. After
washing, the percentage of fluorescent
cells (cell-to-bead ratio of 1:5 or 1:25)
was determined using flow cytometry.
Results are presented as the mean relative binding (HSA-coated beads (1:5)
is set at 1) ⫾ SEM from three independent experiments (each experiment
was performed in triplicate). ⴱ, p ⬍
0.05 and ⴱⴱ, p ⬍ 0.001.
FIGURE 2. Mycobacterial ␣-glucan is a ligand for DC-SIGN. A, Differential binding of DC lectin-Fc constructs to purified ␣-glucan. Binding
of MR-Fc, DC-SIGN-Fc, and dectin1-Fc to ␣-glucan was determined by
ELISA in the absence (⫺) or presence
of EDTA, mannan, or DC-SIGN
blocking Ab AZN-D2. B and C, The
interaction of ␣-glucan with cell surface localized DC-SIGN was investigated by incubating mock cells (Raji
alone) and Raji cells expressing DCSIGN (B) or DCs (C) with ␣-glucan or
HSA-coated control beads at a cell-tobead ratio of 1:25 for 30 min at 37°C
in the absence or presence of mannan
or Ab AZN-D2. After washing, the
percentage of fluorescent cells was determined using flow cytometry. Results are presented as the mean binding ⫾ SEM from three independent
experiments (each experiment was
performed in triplicate). ⴱⴱ, p ⬍ 0.001.
abrogated their capability to bind M. bovis BCG at all three multiplicities of infection (MOIs) tested. Overall, these data demonstrate
that DC-SIGN functions as a cellular receptor for M. tuberculosis
␣-glucan. Additionally, they show that the observed binding of ␣-glucan to the DCs, as demonstrated by the ability of AZN-D2 to block
the interaction, was primarily dependent on this lectin.
DC-SIGN recognizes ␣-1,4-glucan polymers independently of
their origin
DC-SIGN is a C-type lectin receptor that recognizes N-linked high
mannose oligosaccharides and branched fucosylated structures
(40 – 42). Cocrystallization experiments of DC-SIGN with mannose structures have shown that DC-SIGN binding is mediated by
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(38, 39). The interaction with DC-SIGN-Fc was specific, as mannan, EDTA, and the blocking anti-DC-SIGN Ab AZN-D2 fully
abrogated binding (Fig. 2A). We also investigated whether ␣-glucan could interact with native DC-SIGN. By incubating ␣-glucancoated beads together with Raji cells (mock) or Raji cells expressing DC-SIGN (Fig. 2B) or with DCs (Fig. 2C), we found that the
␣-glucan-coated beads showed a significantly increased binding as
compared with the HSA-coated control beads. This interaction was
DC-SIGN-dependent, as preincubation of the cells with mannan or
AZN-D2 decreased the binding to the level of the HSA controls.
Finally, we determined whether ␣-glucan could inhibit Mycobacterium-DC-SIGN interactions. As shown in Fig. 3, preincubation
of DCs or Raji cells expressing DC-SIGN with ␣-glucan strongly
The Journal of Immunology
the interaction of a Ca2⫹ with the vicinal, equatorial 3- and 4-OH
groups of internal mannosyl residues (40). This preference for internal residues is unusual since most mannose-binding lectins (e.g.,
the MR) recognize terminal residues (43, 44). Although earlier
studies already demonstrated that DC-SIGN, besides binding to
mannose and fucose, can also interact with glucose and ␣1,4-diglucosyl (maltose) structures, the potential physiological importance of this interaction was not appreciated (45, 46). To determine
FIGURE 4. Interaction of DC-SIGN with ␣-glucan
is mediated by internal glucosyl residues. A, Binding of
DC-SIGN-Fc to HSA (negative control), purified ␣-glucan, bovine glycogen, and enzymatically synthesized
glycogen (␣1-ESG-A) was determined by ELISA. B,
The ability of DC-SIGN to interact with various neoglycoconjugates was assessed by incubating cells (mock
cells or Raji cells expressing DC-SIGN) with beads
coated with biotin-labeled polyacrylamide neoglycoconjugates harboring single glucosyl molecules (Glc1),
␣-(134)-di-glucosyl molecules (Glc2), or ␣-(134)-tetra-glucosyl molecules (Glc4) at a cell-to-bead ratio of
1:10 for 30 min at 37°C. Beads coated with (man)3ara
(Man3) or (ara)6 (Ara6) glycoconjugates served as positive and negative controls, respectively. After washing,
the percentage of cells positive for fluorescence was determined using flow cytometry. Results are presented as
the mean binding ⫾ SEM from three independent experiments (each experiment was performed in triplicate). ⴱⴱ, p ⬍ 0.001 differences as compared with the
HSA control (A) or the beads coated with the Ara6 and
Glc1 glycoconjugates (B).
whether the interaction of DC-SIGN with ␣1,4-glucan was specific
for the mycobacterial-derived compound or represented a general
phenomenon, binding of DC-SIGN-Fc to glycogen, which is
chemically similar to ␣-glucan (14, 15), was tested. As for mycobacterial ␣-glucan, DC-SIGN-Fc strongly interacted with both
sources of glycogen tested, that is, bovine-derived glycogen and
enzymatically synthesized glycogen, thus suggesting that recognition of ␣1,4-glucans by DC-SIGN was a general phenomenon
(Fig. 4A). To determine the minimal structure that was needed for
the interaction, fluorescent streptavidin-coupled beads, coated with
neoglycoconjugates consisting of biotinylated, polyacrylamide
carriers harboring single glucosyl, ␣-(134)-di-glucosyl (maltose),
or ␣-(134)-tetra-glucosyl (maltotetraose) side chains, were tested
for the ability to bind to Raji cells expressing DC-SIGN. Beads
coated with (man)3ara or (ara)6 (47, 48) were used as positive and
negative controls, respectively. As shown in Fig. 4B, all beads
exhibited a similar binding to mock cells (Raji alone). However,
with the Raji cells expressing DC-SIGN, beads harboring (man)3ara
and di- and tetra-glucosyl conjugates displayed strong binding,
whereas those coated with mono-glucosyl- or (ara)6-containing neoglycoconjugates did not. These data demonstrate that the ␣-(134)di-glucosyl unit represented the minimal structure that was recognized
by DC-SIGN. This suggests that, reminiscent of high-mannose-DCSIGN interactions, DC-SIGN probably recognizes ␣-glucans through
the interaction with internal glucosyl residues.
␣-Glucan induces IL-10 production in LPS-primed DCs in a
DC-SIGN-dependent manner
Binding of mycobacterial ManLAM by DC-SIGN induces the production of IL-10 in LPS-primed DCs (37, 39). To determine
whether ␣-glucan was capable of inducing similar effects, DCs
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FIGURE 3. Mycobacterial ␣-glucan inhibits the interaction between
DC-SIGN and M. bovis BCG. Raji cells expressing DC-SIGN and DCs
were incubated with M. bovis BCG expressing dsRed at a MOI of 0.5, 2,
or 8 for 30 min at 37°C in the absence (filled bars (⫺)) or presence (white
bars (⫹)) of 50 ␮g/ml ␣-glucan. The percentage of cells binding M. bovis
BCG was determined using flow cytometry. Results are presented as the
mean binding ⫾ SEM from three independent experiments (each experiment was performed in triplicate). ⴱ, p ⬍ 0.05 and ⴱⴱ, p ⬍ 0.001.
5225
5226
MYCOBACTERIAL ␣-GLUCAN: A NOVEL LIGAND FOR DC-SIGN
were incubated with LPS in the absence or presence of ␣-glucan or
with ␣-glucan alone. As shown in Fig. 5A, stimulation with LPS in
the presence of ␣-glucan significantly increased the relative IL-10
production (⬃4.2-fold) as compared with stimulation with LPS
alone. ␣-Glucan by itself did not induce detectable levels of IL-10.
To investigate whether the effect was specific of IL-10 or also
altered the IL-12/IL-23 axis, the relative amount of IL-12p40 was
also determined. As shown in Fig. 5B, stimulation with LPS in the
presence of ␣-glucan did not significantly alter relative IL-12p40
secretion. Similar results were obtained with ␣-glucan associated
to polystyrene beads (data not shown). Taken together, these results demonstrate that ␣-glucan, like ManLAM, induced the production of IL-10 in LPS-primed DCs.
Recently, the molecular signaling pathway underlying DCSIGN-dependent IL-10 induction in LPS-primed DCs was identi-
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FIGURE 5. ␣-Glucan induces IL-10
production in LPS-primed DCs in a DCSIGN-dependent manner. DCs were
stimulated with ␣-glucan (20 ␮g/ml)
in the absence or presence of LPS (20
ng/ml) or with LPS alone. After 24 h,
the supernatants were harvested and
the relative IL-10 (A) and IL-12p40
(B) protein levels were determined.
Additionally, the cells were stimulated
as in A in the absence or presence of
AZN-D2 or anacardic acid (compounds were added 2 h before stimulation), after which the relative IL-10
production (C) and IL-10 mRNA expression (D) were determined using
ELISA and quantitative real-time PCR
analysis, respectively. In all cases, the
values obtained for LPS-stimulated
cells were set at 1. Results are presented as the mean ⫾ SEM from at
least five independent experiments
(each experiment was performed in
triplicate). ⴱ, p ⬍ 0.05 and ⴱⴱ, p ⬍
0.001; n.s., nonsignificant. E, Specific
p65 phosphorylation (Ser276; black
bars) and acetylation (Lys310; gray
bars) was determined by ELISA on
nuclear extracts of DCs stimulated for
1 h with ␣-glucan or ManLAM (20
␮g/ml) in the absence or presence of
LPS (20 ng/ml) or anacardic acid.
Data represent the means ⫾ SEM of
two independent experiments.
fied (37). It was demonstrated that ligand binding by DC-SIGN
activates the serine/threonine kinase Raf-1, which leads to phosphorylation and acetylation of the NF-␬B subunit p65 and, consequently, a prolonged and increased transcription of IL-10 (37). To
investigate the DC-SIGN dependency of ␣-glucan-induced IL-10
production, DCs were pretreated with two specific inhibitors, after
which the consequences for ␣-glucan-dependent IL-10 induction
were analyzed. Preincubation with the blocking Ab AZN-D2 abrogated the ␣-glucan-induced secretion of IL-10 (Fig. 5C). This
inhibition was also observed at the mRNA level, as pretreatment
with the Ab significantly reduced the expression of IL-10 mRNA
(Fig. 5D). To test whether the induction was also dependent on the
acetylation of the p65 subunit of NF-␬B, the DCs were preincubated with anacardic acid. This CREB-binding protein/p300-specific histone acyltransferase inhibitor prevents p65 acetylation and
The Journal of Immunology
5227
was previously shown to block ManLAM-induced expression of
IL-10 (37). Consistent with this earlier report, pretreatment with
anacardic acid blocked the ␣-glucan-induced production of IL-10
(Fig. 5C). This inhibition was also apparent at the mRNA level, as
the addition of anacardic acid significantly reduced the amount of
expressed IL-10 (Fig. 5D). Furthermore, analysis of the phosphorylation and acetylation status of p65 extracted from the nucleus of
stimulated DCs demonstrated that ␣-glucan, like ManLAM, induced the phosphorylation of p65 serine residue 276 and the acetylation of lysine residue 310, with this latter process being inhibited by anacardic acid (Fig. 5E). Overall, these data demonstrate
that the ␣-glucan-induced IL-10 production in LPS-primed DCs,
as for ManLAM, was dependent on DC-SIGN and acetylation of
the p65 subunit of NF-␬B.
Mycobacterial ␣-glucan is not a ligand for TLR2
FIGURE 6. TLR2 activation by purified ␣-glucan is caused by contamination with lipopeptides. A, HEK293 cells transfected with TLR2 were
stimulated with increasing amounts of ␣-glucan, with LPS (50 ng/ml), with
Pam3CysK4 (50 ng/ml), or with H2O (⫺) as a negative control for 24 h at
37°C. Following stimulation, the supernatants were harvested and analyzed
for IL-8. B, HEK293 TLR2 cells were stimulated with ␣-glucan (50 ␮g/ml)
that was pretreated or not with amyloglucosidase, 1% hydrogen peroxide
(H2O2), or phenol. Stimulation with hydrogen peroxide-treated
Pam3CysK4 (50 ng/ml) served as a positive control for lipopeptide inactivation. In both A and B, the results represent the mean IL-8 production ⫾
SEM from three independent experiments (each experiment was performed
in triplicate). ⴱⴱ, p ⬍ 0.001; n.s., nonsignificant. C, Consequence of amyloglucosidase, hydrogen peroxide, and phenol reatment on the integrity of
␣-glucan was assed by monitoring the reactivity of an anti-␣-glucan mAb
toward the (pretreated) ␣-glucan. The figure shows the reactivity of the Ab
toward a 5-fold serial dilution series of (pretreated) ␣-glucan (starting at 20
␮g/ml).
any cytokine secretion by itself (Fig. 7B). These results are consistent with the earlier report that ManLAM alone does not induce
cytokine secretion by DCs (37). Finally, we tested whether the
phenol-treated ␣-glucan could still block the interaction between
DCs and M. bovis BCG. Similar to the results obtained for untreated ␣-glucan (Fig. 3), phenol-treated ␣-glucan efficiently
blocked the interaction between M. bovis BCG and the DCs (Fig.
7C), thus ruling out that the block was caused by the lipopeptide
contamination.
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Extracellular ␣-glucans are not unique to mycobacteria, and similar molecules can be found in a variety of species. Some of these
␣-glucans have been shown to possess immune-stimulating properties, for which the underlying mechanisms remain largely unknown (33, 49 –51). Interestingly, for the fungus Pseudallescheria
boydii, it was recently reported that the stimulating activity of its
␣-glucan was dependent on the activation of TLR2 (51). This report, together with the earlier observation that mycobacterial
␣-glucan alone induced low, but detectable amounts of IL-12p40
(Fig. 5B), prompted us to investigate whether the mycobacterial
compound may also be a ligand for TLR2. To test this, TLR2transfected HEK293 cells were incubated with serial dilutions of
purified ␣-glucan, after which IL-8 production was analyzed. As
shown in Fig. 6, stimulation with mycobacterial ␣-glucan activated
TLR2 in a dose-dependent manner (Fig. 6A). Consistent with the
TLR2 dependency, the cells were also activated by the TLR2 agonist Pam3CysK4, but not by the TLR4 agonist LPS (Fig. 6A).
Besides ␣-glucan, many structurally unrelated compounds, including various glycolipids, lipopeptides, proteins, and polysaccharides, have been shown to induce TLR2 activation (25, 52). However, despite the sometimes convincing evidence, the activity of
many of these compounds was later on shown to be caused by
contamination with lipopeptides (53–55). To investigate the possibility that the observed TLR2-stimulating activity of ␣-glucan
was caused by lipopeptide contamination, ␣-glucan was pretreated
with hydrogen peroxide, phenol, and amyloglucosidase and retested for the activity on HEK293 TLR2 cells. As shown in Fig.
6B, treatment of ␣-glucan with hydrogen peroxide and phenol,
treatments that inactivate (25) and remove (56) lipopeptides, respectively, completely abrogated the TLR2-stimulating activity.
However, activity was recovered in the phenol extract (data not
shown). In contrast, pretreatment with amyloglucosidase, which
completely degraded the ␣-glucan (Fig. 6C), did not significantly
reduce the TLR2-mediated activity (Fig. 6B). These data demonstrate that the observed TLR2-stimulating activity was not mediated by the ␣-glucan but was most likely due to a low-level contamination with lipopeptides. This conclusion was further
supported by the observation that various types of glycogen, both
natural and synthetic, were unable to activate TLR2 (data not
shown). To investigate whether the removal of lipopeptides would
alter the conclusions drawn from our earlier experiments, DCs
were stimulated with phenol-treated ␣-glucan in the absence or
presence of LPS. As shown in Fig. 7, phenol-treated ␣-glucan, as
for the untreated ␣-glucan (Fig. 5A), specifically induced the production of IL-10, confirming the dependency on DC-SIGN (Fig.
7A). However, in contrast to the earlier experiment in which untreated ␣-glucan was found to induce low, but detectable amounts
of IL-12p40 (Fig. 5B), the phenol-treated ␣-glucan did not induce
5228
MYCOBACTERIAL ␣-GLUCAN: A NOVEL LIGAND FOR DC-SIGN
Discussion
Polysaccharide capsules are expressed by many pathogenic bacteria and play an important role during infection (10, 11). Although
the presence of a capsule on mycobacteria was already recognized
many decades ago (57), so far only a few studies have been aimed
at investigating its function. One important requirement for obtaining insight into its functional role is to identify host receptors
that recognize its constituents. The polysaccharide composition of
the capsule is conserved among mycobacteria and predominantly
consists of an ␣-(134)-glucosyl polymer known as ␣-glucan (up
to ⬃80% of the capsular polysaccharide content in M. tuberculosis) (12, 13). So far, only one potential host ␣-glucan receptor has
been identified. Cywes and colleagues demonstrated that both mechanical and enzymatic removal of the capsule, and in particular
that of the capsular ␣-glucan, abolished the nonopsonic binding of
M. tuberculosis to Chinese hamster ovary cells expressing CR3
(18). Additionally, it was shown that capsular ␣-glucan was able to
inhibit Mycobacterium-CR3 interactions. These findings are puzzling, as CR3 has been shown to bind well to ␤-glycans but not to
␣-glycans (58). Nevertheless, these data suggest that CR3 acts as
a cellular ␣-glucan receptor. More recently, in a study performed
by Gagliardi and coworkers, it was shown that incubation of
monocytes with capsular ␣-glucan before differentiation blocks
CD1 expression on the monocyte-derived DCs and suppresses
their IL-12 production (22). Although these immunosuppressive
effects could be ascribed to the glucan, the host receptors involved
were not identified, nor were the signaling routes explained. Therefore, our aim was to identify (additional) host ␣-glucan receptors
and investigate their role in the Mycobacterium-host interaction.
To do this, we first performed a screen in which different types
of primary immune cells were checked for their ability to interact
with ␣-glucan. This experiment indicated that ␣-glucan bound to
macrophages and, interestingly, also to DCs (Fig. 1). DCs are important immune cells that are pivotal for the induction of an adap-
tive immune response. This characteristic makes DCs an attractive
target for pathogenic microbes, as illustrated by the large number
of pathogens, including pathogenic mycobacteria, that modulate
their function (59 – 61). For this reason, together with the observation that ␣-glucan binding to the macrophages was generally
weak, we focused on the DC-␣-glucan interaction. Several DC
lectins were previously shown to interact with mycobacteria.
These include the MR (62), DC-SIGN (38, 39), and dectin-1 (63,
64). By using Fc constructs of these lectins, we were able to show
that ␣-glucan was specifically recognized by DC-SIGN (Fig. 2A).
Experiments with native DC-SIGN confirmed these results and
demonstrated that ␣-glucan represented a bona fide ligand for this
lectin (Fig. 2, B and C). Furthermore, by using a blocking Ab, we
could demonstrate that ␣-glucan binding to DCs was primarily
dependent on DC-SIGN (Fig. 2C). The inability of dectin-1-Fc to
bind ␣-glucan was expected, as this receptor is mainly involved in
the recognition of ␤-linked glucans (65). However, the MR, like
DC-SIGN, harbors an EPN (in one-letter amino acid code) motif
and has been shown to recognize mannose, fucose, N-acetylglucosamine, and glucose-containing structures (66 – 68). However, in
contrast to DC-SIGN, which recognizes internal glycosyl residues,
the MR interacts with terminal moieties (43, 44). This characteristic may explain the differential recognition, as the relative
amount of nonreducing termini in ␣-glucan is expected to be low
as compared with the number of internal motifs. This view is supported by the observation that the MR-Fc construct showed a low,
but significant binding to glycogen (data not shown). As glycogen
possesses a more open structure and shorter backbones than does
␣-glucan (16), it is expected to express a higher number of accessible terminal glucosyl residues. The assumption that ␣-glucan recognition by DC-SIGN is potentially mediated by internal glucosyl
residues was further supported by the observation that beads
coated with ␣-(134)-di-glucosyl (maltose) moieties (in the form
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FIGURE 7. Phenol-treated ␣-glucan induces IL-10
production in LPS-primed DCs and inhibits the interaction between DC-SIGN and M. bovis BCG. DCs were
stimulated with phenol-treated ␣-glucan in the absence
or presence of LPS or with LPS alone. After 24 h, the
supernatants were harvested and the relative IL-10 (A)
and IL-12p40 (B) protein levels were determined. C,
DCs were incubated with M. bovis BCG expressing
dsRed at a MOI of 0.5, 2, or 8 for 30 min at 37°C in the
absence (filled bars (⫺)) or presence (open bars (⫹)) of
50 ␮g/ml phenol-treated ␣-glucan. The percentage of
cells binding M. bovis BCG was determined using flow
cytometry. In all cases, results are presented as the
means ⫾ SEM from three independent experiments
(each experiment was performed in triplicate). ⴱⴱ, p ⬍
0.001; n.s., nonsignificant.
The Journal of Immunology
matic effect on the biological activity. Additionally, simply converting the thioether linkage present in lipopeptides into a sulfoxide (by hydrogen peroxide treatment) completely abrogated the
activity (26). This tight relation between structure and function can
be understood from x-ray data of a TLR1-TLR2-lipopetide complex (74). Hydrophobic pockets on the surface of the TLRs will, to
fit in, put strong restrictions on the nature and spatial distribution
of the ligand acyl chains. We therefore investigated the possibility
that the TLR2 activity of capsular glucan was caused by contaminating lipopeptides by two independent procedures: removal of
lipopeptides by phenol extraction, and inactivation of lipopeptides
by treatment with hydrogen peroxide. As shown in Fig. 6B, pretreatment of ␣-glucan with both hydrogen peroxide and phenol
extraction fully abrogated its TLR2-activating potency. In contrast,
the activity after degradation with amyloglucosidase was sustained. These results demonstrate that the TLR2 activation was not
mediated by ␣-glucan but almost certainly was caused by contaminating lipopeptides. These copurified lipopeptides were present at
very low concentrations, as they remained undetected by 1H NMR
analysis after ␣-glucan was purified. Fig. 6 shows that the presence
of 0.1% of lipopetides in the ␣-glucan fraction (a level that cannot
be detected by NMR) would be sufficient to account for the activity
observed. Our case is by no means novel, and the biomedical literature shows that low-level endotoxin and/or lipopeptide contamination, undetectable with chemical methods, has been haunting
scientists for many years (25). Overall, our findings demonstrate
that the analysis of putative TLR2 ligands, especially those that do
not harbor hydrophobic domains, should be performed with great
care, and that suitable controls, for example, treatment with phenol
and hydrogen peroxide, should be included. Furthermore, it is clear
that before mycobacterial ␣-glucan is used in biological assays, an
additional purification step that removes lipopeptides, for example,
treatment with phenol, should be performed.
Here we have demonstrated that mycobacterial ␣-glucan represents a bona fide ligand for DC-SIGN. This has added a second host ␣-glucan receptor and has broadened the potential target cell population to also include DCs. This observation,
together with the notion that mycobacteria shed high amounts
of ␣-glucan (12), suggests that the ␣-glucan capsule may fulfill
an important role in pathogenesis. To investigate this issue, the
generation of ␣-glucan-deficient mutants will be of major benefit. However, despite great endeavor, the construction of such
mutants has currently been unsuccessful. Recently, it was
shown that mutation of Rv3032, a homolog of the glycogen
synthase glgA, greatly reduced the synthesis of methyl glucose
LPS and cytosolic glycogen, whereas the levels of extracellular
␣-glucan remained unchanged (75). However, deletion of a second glgA homolog, that is, Rv1212c, resulted in reduced
amounts of ␣-glucan with no effect on the levels of methyl
glucose LPS and glycogen. Still, importantly, the Rv1212c mutant could be fully complemented by overexpression of Rv3032.
Furthermore, a double mutant of Rv1212c and Rv3032 could not
be generated, indicating that the presence of at least one of the
copies was required for viability (75). These findings suggest
that ␣-glucan and glycogen may share, at least in part, a common biosynthetic pathway. However, as the two polysaccharides also exhibit important structural differences (16), discrepancies between the biosynthetic pathways must exist.
Nevertheless, the observation that enzymes involved in the biosynthesis of ␣-glucan are essential for viability warrants its biosynthetic pathway as an interesting target for TB drug
development.
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of side chains of polymeric neoglycoconjugates) strongly interacted with DC-SIGN, whereas those coated with mono-glucosyl
residues did not (Fig. 4B). Also in a previous study, maltose was
shown to be a much more effective competitor than glucose (Ki of
0.27 and 8.08 mM, respectively) for inhibiting interactions between gp120 and DC-SIGN (46). However, the conclusion that
DC-SIGN binds ␣-glucan through internal glucosyl residues also
poses a fundamental problem in that it is known to preferentially
recognize equatorial 4-OH groups (40). As ␣-glucan is composed
of ␣-(134)-linked residues, the internal 4-OH groups are not
available for binding to DC-SIGN. This raises the intriguing question of how ␣-glucan is recognized, and additional experiments
will be needed to provide an answer to this important question.
Previously, ligand binding by DC-SIGN was shown to induce
IL-10 production in LPS-primed DCs. This phenomenon was first
observed for ManLAM, an abundant mycobacterial cell wall glycolipid (39). It was proposed that mycobacteria target DC-SIGN to
induce IL-10 production and thereby subvert the host immune response (39). Consistent with this result, we observed that also
␣-glucan induced IL-10 production in LPS-primed DCs (Fig. 5A).
This suggests that ␣-glucan, as for ManLAM, may promote immune suppression. However, one potential flaw in this reasoning is
that all of the experiments showing IL-10 induction by mycobacterial compounds, including our own, were performed in the context of TLR costimulation (TLR4 in particular), the role of which
in M. tuberculosis infection remains unclear (69). Elucidation of
the molecular pathways underlying DC-SIGN-TLR cross-modulation has only just begun. Important progress into this field was
made when it was demonstrated that DC-SIGN ligation activates
the serine/threonine kinase Raf-1, leading to the phosphorylation
and acetylation of the NF-␬B subunit p65 and a prolonged and
increased transcription of IL-10 (37). However, it remains unclear
whether this route is the only way by which DC-SIGN influences
TLR signaling or that alternative regulatory mechanisms may exist. Importantly, stimulation of DC-SIGN alone does not seem to
induce IL-10 production (70). Additionally, in the bronchoalveolar
lavage of patients with TB, IL-10 could not be detected (71). Furthermore, the cross-talk between DC-SIGN and TLR2—the TLR
that is probably most relevant in the context of mycobacterial infections— has not been investigated. Nevertheless, the potential
importance of DC-SIGN ligation during infection is illustrated by
the large number of DC-SIGN ligands that (pathogenic) mycobacteria may express. Together with ManLAM (39), lipomannan (72),
mannose-capped arabinomannan (72), two mannosylated glycoproteins (72), and the phosphatidylinositol mannosides (73), ␣-glucan represents the seventh documented mycobacterial ligand for
DC-SIGN. Interestingly, it has been shown that in patients with TB
up to 70% of alveolar macrophages show DC-SIGN expression
(71). It was demonstrated that infection with M. tuberculosis induced DC-SIGN expression, both on infected and noninfected
macrophages, and that the DC-SIGN-expressing cells were more
prone to infection than were their DC-SIGN-negative counterparts
(71). Overall, these findings strongly suggest that DC-SIGN ligation is an important process during mycobacterial infection. However, how M. tuberculosis exactly benefits from this interaction
remains unclear.
To date, a wide diversity of structurally unrelated molecules
have been claimed to induce TLR2 activation. These structures
include various (lipo-)proteins and lipopeptides, glycolipids, peptidoglycans, and other polysaccharides and even whole viruses (for
a comprehensive review, see Ref. 25). This apparent promiscuity
contrasts strongly with data suggesting a very high specificity (26).
In the latter study, it was shown that variations in the acylation of
a (synthetic) lipopeptide, an established TLR2 ligand, had a dra-
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Acknowledgments
We thank Drs. T. Geijtenbeek and S. I. Gringhuis (Academic Medical
Center, University of Amsterdam, Amsterdam, The Netherlands) for providing DC-SIGN-Fc, mAb AZN-D2, DC-SIGN-transfected Raji cells, and
technical advise and assistance; Dr. G. Brown (University of Cape Town,
Cape Town, South Africa) for providing dectin-1-Fc; and Dr. L. MartinezPomares (University of Nottingham, Nottingham, United Kingdom) and
Dr. R. Stillion (University of Oxford, Oxford, United Kingdom) for providing MR-Fc. Furthermore, we thank Dr. D. Golenbock (University of
Massachusetts Medical School, Worcester, MA) for providing the HEK293
TLR2 cell line. We also thank Dr. U. Zähringer (Leibnitz Research Center,
Borstel, Germany) for crucial discussions on TLR2 ligands.
Disclosures
23.
24.
25.
26.
27.
28.
The authors have no financial conflicts of interest.
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